Characterization of chitin and chitosan derived from Hermetia illucens, a further step in a circular economy process

Composition of raw samples

Results regarding composition of larvae, pupal exuviae and dead adults of H. illucens related to the dry mass of the samples are reported in Table 1.

The three samples had a significantly different composition. Larvae had the greatest amounts of lipids (23.0%), adults the highest protein content (49.0%), while minerals (16.0%) and fibres (53.3%) were higher in pupal exuviae. Pupal exuviae were also the insect sample with the highest chitin content (25.5%), representing the biomass of choice for the production of chitin and its derivative, chitosan. According to literature, whole insects contain 30–60% protein, 10–25% lipid, 5–25% chitin and 2–10% minerals^{31,58,59,60,61}. All these components are reported in wide ranges, as the composition of insects varies, especially depending on their species and stage of development. Our data regarding larvae, pupal exuviae and dead adults from the H. illucens are within these ranges. Similarities were also found in the average composition of the different developmental stages of H. illucens reported by other authors^62,63,64,65, except for lipids. Indeed, our samples had a lower lipid content than those reported for the same insect biomass (3–40%, 8 and 27% for larvae, exuviae and adults, respectively)^62,63. These differences could be due to the different feeding substrate of larvae and to the different methodology used for the analysis of lipid content.

Chitin purification from H. illucens

Chitin was purified from H. illucens larvae, pupal exuviae and dead adults (Fig. 1a), leaving a part of each chitin sample as it was, unbleached (Fig. 1a(A–B–C)), and another bleached (Fig. 1a(D–E–F)). Bleaching treatment resulted in a chitin with a whiteness similar to commercial chitin and, even brighter, in case of adults bleached chitin (Fig. 1a(F)). A qualitative assessment of the insect chitin purification process is generally reported in the literature, instead, data providing a comprehensive quantitative assessment is lacking. Even less data is available on the extraction process of chitin from H. illucens. Results of the chitin extraction process were expressed in terms of efficiency of the purification method applied, chitin recovery during it, purity and yield of the obtained chitin.

Biomass recovery and chitin yield

Biomass recovery from the insect samples after each purification step was calculated for all samples (Table 2). Due to very few numerical data providing this value, our results can only be compared with values reported by Khayrova et al.^57,65.

The highest recovery was obtained with pupal exuviae during all extraction procedures, on the other hand adults had the lowest. Specifically, biomass recovery after demineralization (DM) of both larvae and pupal exuviae was similar to or slightly higher than that obtained by Khayrova et al.^57,65 (58 and 74%, respectively), in contrast to that from adults which was lower (68% vs 87%)⁶⁵. After deproteinization (DP), lower amounts were recovered from demineralized chitin than values reported by Khayrova et al.^57,65 (46, 77 and 24% for larvae, exuviae and adults, respectively). Values reported by Hahn et al.⁶⁶, referring to larval exoskeletons of H. illucens (79% after DM and 34% after DP) were close to those for pupal exuviae. These results revealed correlations between protein content of the starting sample and the chitin recovery after the DP step. Indeed, the adults, the sample with the highest starting protein content (49%), had the lowest percentage of chitin recovery after DP (14%); in contrast, pupal exuviae, with the lowest starting protein (30%), gave the highest percentage of recovery (41%). In accordance with the composition of raw samples of H. illucens (Table 1), adults contained, similarly to larvae, a higher protein and lipid content (about 70%) than that reported for pupal exuviae (about 35%), resulting in a lower final chitin content. These compositional differences can be explained considering that larvae and adults catabolize lipids and proteins to produce the energy necessary for growth and reproduction, respectively; pupal exuviae on the other hand, represent a waste product resulting from the moulting process and are mainly composed of chitin.

Yields of unbleached chitin from larvae, pupal exuviae, and adults were 13, 31, and 9%, respectively (Table 2). After bleaching, these yields decreased slightly to 10, 23, and 6%, but no significant differences were found between the two values for each sample (larvae Χ² = 0.2, p = 0.66; pupal exuviae Χ² = 1.24, p = 0.26; adults Χ² = 0.29, p = 0.59), thus demonstrating the bleaching treatment did not affect chitin yield but favored its degree of purity. Chitin yields are within or higher than the average range reported for chitin from insects (5–15%)³² and chitin from crustaceans (5–30%)^20,67,68,69. For insects, the highest values, between 31 and 36%, were obtained from larval exoskeletons of H. illucens⁶⁶, cicada sloughs⁷⁰ and adults of Apis mellifera⁷¹. The yields of bleached chitin, obtained in the present work, were comparable to or higher than those obtained by other authors using the same developmental stage of H. illucens^{63,64,72,73,74}. In a similar range (6–26%) are the yield values obtained by Zhou et al.⁷⁵ using natural deep eutectic solvents for the purification of chitin from H. illucens prepupae. It should be considered that the yield of chitin can vary greatly depending on the species, the developmental stages, the body parts, and the extraction methods used.

Efficiency of the purification process

Composition of larvae, pupal exuviae and dead adults after each extraction step, as well as composition of the final chitin, were determined and presented in Table 3.

Minerals were significantly reduced by DM treatment with formic acid for all insect samples (larvae Χ² = 6.3, p = 0.01; pupal exuviae Χ² = 9.5, p = 0.002; adults Χ² = 4.1, p = 0.04), to remain constant during the following steps, with the only exception of pupal exuviae, where bleaching induced a further decrease. It resulted in 82, 85, and 87% DM efficiency for larvae, pupal exuviae, and adults, respectively. Only Hahn et al.⁶⁶ used formic acid for DM of H. illucens larval exoskeletons, removing 84% of minerals. A similar efficiency (84.1%) was achieved by Khayrova et al.⁶⁵ using hydrochloric acid on H. illucens pupal exuviae, while only 47.5% minerals was removed from adult flies applying the same acid. On H. illucens prepupae, on the other hand, the use of natural deep eutectic solvents, achieved a DM efficiency of 98%⁷⁵. Our results proved that the use of an organic acid, with less environmental impact and less potentially negative effect on the final chitin⁷⁶, can therefore remove minerals from different H. illucens samples with similar or higher efficiency than a mineral acid. This was also demonstrated by other studies on insects and crustaceans: oxalic acid removed a higher percentage of minerals (85%) from house crickets⁷⁷, compared to that obtained using hydrochloric acid, under the same conditions on both flies (39.5%)⁷⁸ and two-spotted crickets (8–21%)⁷⁹, lactic and acetic acid were used for DM on shrimp shells with comparable efficiency to that obtained with hydrochloric acid^76,80.

Proteins, on the other hand, were decreased significantly by DP treatment with sodium hydroxide in all insect samples (larvae Χ² = 37.9, p < 0.001; pupal exuviae Χ² = 26.2, p < 0.001; adults Χ² = 57.3, p < 0.001) (94, 92 and 97% efficiency), to remain constant until the end of the purification process; in larvae only, a significant reduction of the protein content was already found post DM. It resulted in 94, 92 and 97% DP efficiency for larvae, pupal exuviae, and dead adults, respectively. A slightly lower value (86–87%) was obtained with similar reaction conditions on Musca domestica pupae and Gryllus bimaculatus adults^78,79. A higher value (97%) was obtained using natural deep eutectic solvents on H. illucens prepupae⁷⁵.

Purity of chitin extracted from the three biomasses of H. illucens was expressed as chitin content. The chitin content increased as the extraction process advanced in all insect samples, with the major rise recorded after the DP where unbleached chitin is obtained. The unbleached chitins were very similar to each other (chitin content amounting to 73.4, 76.9 and 77.8%, respectively), except for the residual mineral and protein content, which was lower in adults than in the other ones. The bleaching treatment did not significantly affect the acid detergent fibre (ADF) value and, consequently, the final content of the bleached chitin, amounting to 84, 86.8 and 85.3% for larvae, pupal exuviae and dead adults, respectively. These values highlighted the suitability of the applied purification method for the production of a bleached chitin with a degree of purity similar to the commercially available polymer (88.1%).

Degree of purity of bleached chitin extracted from different insect species mostly ranges from 85 to 97%^{64,66,75,81,82}. Given the same insect biomass, the observed differences in chitin purity may be due to the different purification methods applied, in terms of reagents, concentrations, and reaction times, as well as the different methods used to calculate this degree^{64,66,75,81,82}.

Chitosan production

Chitosan was produced by heterogeneous deacetylation of both unbleached and bleached chitin extracted from the three biomasses of H. illucens, thus obtaining six different chitosan samples (Fig. 1b). As expected, chitosan produced from unbleached (Fig. 2b(A–B–C)) chitins were darker than that from bleached chitins (Fig. 2b(D–E–F)), especially adults unbleached chitosan (Fig. 2b(C)). All bleached chitosan also appeared darker than their respective bleached chitins, especially the chitosan of dead adults and pupal exuviae. This browning is probably due to the high temperatures used for the deacetylation reaction, inducing some saccharide dehydration leading to double bonds formation^83,84.

Chitosan recovery and chitosan yield

Yields related to chitin (chitosan recovery after deacetylation) and to the original insect sample were determined for all chitosan samples and are presented in Table 4. For all insect samples, the yield of bleached chitosan related to chitin was higher than the yield of the respective unbleached one (range 25–28%), with the highest values obtained for chitosan from pupal exuviae and adults (42 and 41%, respectively). These significantly different chitosan yield values suggest how, for the same conditions used, the bleaching or not of the chitin influences its deacetylation capacity. Indeed, the unbleached sample presents catechol compounds that, cross-linked to the α-chitin chains, probably hide the acetyl groups and limit the access by NaOH molecules. The reaction parameters should be regulated by trying to force more the deacetylation reaction on the unbleached chitin samples. The yield of chitosan related to the original insect biomass followed the same trend, but with a smaller gap between bleached and unbleached chitosan for each sample. These yield values did not appear to be affected by the bleaching treatment. Yield of chitosan derived from H. illucens by heterogeneous deacetylation was reported only by Khayrova et al.⁵⁷ for larvae (53% related to chitin) and Hahn et al.⁶⁶ for larval exoskeletons (47% related to chitin and 16% related to the initial biomass). The yields obtained in the present work are in the range of chitosan produced from insects (2–8%)³² and are slightly lower than those of crustacean-derived chitosan (4–15%)^{67,68,69,85,86}. This difference can be explained considering that insect biomass has a higher protein and lipid content than crustaceans, which may lower the final polymer yield⁸⁷. However, as reported for chitin, chitosan yield can be affected by various factors, including the source, the purification methods and the deacetylation treatments applied to chitin⁸⁸.

Chitin and chitosan characterization

Fourier-transformed infrared spectroscopy (FTIR)

Spectra resulting from FTIR analysis of both unbleached and bleached chitins from H. illucens larvae, pupal exuviae and dead adults are shown in Fig. 2a(A,B), in comparison to the commercial crustacean-derived chitin. All the characteristic peaks of chitin were detected in all samples at their specific wavelengths: 1310–1320 cm⁻¹ (CN-stretching, amide III), 1550–1560 cm⁻¹ (NH-bending, amide II), 1650–1655 cm⁻¹ (CO-stretching, amide I), 3100–3110 cm⁻¹ (NH-symmetric stretching), 3255–3270 cm⁻¹ (NH-asymmetric stretching) and 3430–3450 cm⁻¹ (OH-stretching). The α-form was confirmed for all chitin samples produced from H. illucens by observing the two Amide I band splits, around 1620 and 1650 cm^{−156,70,89,90,91}. The spectra of all chitins showed a structural similarity with the commercial polymer. No significant differences were observed either among chitin extracted from the different starting insect materials, in accordance to other authors^63,72,73, or for each sample, between the bleached chitin and the respective unbleached one. Some differences in peak wavelengths are probably due to the different natural sources and the extraction process applied. Chitin acetylation degree (AD) was also determined from the spectra (Table 5).

The AD values of all chitin samples are within the range of AD reported for both insect-derived chitin and the commercial one (80 -100%), considering only the AD determined by FTIR^{63,89,92,93,94}. Results of the present work were also similar to those already reported for chitin derived from H. illucens^56,63,64. The AD of chitin obtained from adults was the highest. These comparable values among the samples (larvae, pupal exuviae, dead adults) of H. illucens indicated that AD was not subject to large variations in the growth stages, confirming that reported by other authors for the same insect⁶³ and other species⁸².

Spectra resulting from FTIR analysis of chitosan samples are shown in Fig. 2a(C,D), in comparison with the commercial one. As reported for chitin, characteristic peaks confirming the identity of chitosan were detected, specifically the NH-bending (amide II) and CO-stretching (amide I) bands around 1655 (amide I) and 1590 cm⁻¹ (NH₂ bending), respectively^{86,90,91,95,96}. No significant differences were observed between the spectra of bleached chitosan and the respective unbleached sample for adults only. The N–H and O–H stretching bands in the 3000–3600 cm⁻¹ region are more complex for chitin than for chitosan, in agreement with the presence of different groups (amine or amidic N–H, for instance). Moreover, this region is affected by inter-macromolecular interactions that seem to be more complex in chitin. These interactions are connected to supramolecular organization (presence of hydrogen bonds, formation of crystals, etc.), depending on material morphology. The deacetylation, being a chemical reaction that affects the chitin morphological structure, had reasonably affected intermolecular and supramolecular organization, as deductible by comparing chitin and chitosan spectra.

X-ray diffractometry (XRD)

XRD analysis was performed to determine crystallinity of chitin and chitosan from H. illucens.

The spectra obtained for chitins are shown in Fig. 2b(A,B). Similarly to the commercial sample, all chitins showed the significant sharp peaks at 9° and 19° and the three/four weak peaks around 13°, 21°, 23° and 26°, confirming the α-form of the polymer^70,89,97. No significant differences were found in the spectra between unbleached and bleached chitin; only both chitins derived from pupal exuviae were different for the presence of more intense peaks between 19° and 26° (Fig. 2b(A,B). The peaks of all chitin samples were found to be very similar to those reported for other insect species, in range 9°–26°^89,92,98,99 and for H. illucens itself^{56,64,72,73,97}.

The determination of the CrI from the XRD data revealed significant differences among chitins derived from various developmental stages, comparing them to commercial chitin (Table 5). Generally, all bleached chitins had a slightly lower CrI than the respective unbleached samples (although not significantly). This suggests a possible detrimental effect of the bleaching treatment on the crystalline structure of polymer.

Adults-derived chitin was the most crystalline and not different from commercial one, followed by larvae with slightly lower values, while the lowest CrI were obtained from pupal exuviae derived chitins with values below 70%. Crystallite size was similar among all chitins, including the commercial one (Table 5).

Similar results were also reported by other authors for chitin produced from different biomasses of H. illucens, with adult chitin always being more crystalline than chitin from other stages^56,72,73. It was inferred that the crystallinity of chitin increases gradually, thus showing a more ordered structure, at the life stages of the dipteran, particularly from pupa to adult^56,72.

The CrI values of crustacean and insect chitin fall within a wide range, from 40 to 90%, mainly 60–80%, as they are depending on the source, in terms of species, growth stage and gender, and on the purification process^{32,74,75,96,97,99,100,101}. According to its crystallinity, chitin can be used in different fields: a more amorphous chitin (low CrI) has absorbent properties that make it effective in removing contaminants, such as heavy metals, and therefore useful in water treatment and industrial applications¹⁰²; on the other hand, the high crystallinity of chitin can be a positive aspect for the formulation of chitin nanofibrils, applied in the cosmetic and biomedical field^103,104.

XRD patterns of both unbleached and bleached chitosan are shown in Fig. 2b(C,D).

In the XRD analysis of all chitosan samples, the two main sharp peaks around 10° and 20°, were observed. These peaks were similar to those reported for insect-based and crustacean chitosan^{32,86,89,105,106}. As reported for chitin, no significant differences were found in the spectra between unbleached and bleached chitosan; bleached chitosan derived from larvae and pupal exuviae were different from their respective unbleached samples by the presence of a more and less intense peak at 9°, respectively. CrI values of chitosan samples were all statistically similar, including the commercial one, and ranged from 74 to 86% (Table 4). There is a tendency for bleached samples to be slightly more crystalline than unbleached ones, although not significantly. Due to the lack of studies on the effect of chitin bleaching on the crystallinity of chitosan, it was not possible to compare results of the present work with others in the literature. The crystallinity of chitosan obtained from H. illucens was higher than that reported for the other insects (33–69%)^{86,105,106,107,108} and more similar to that of commercial one. Crystallite size was similar among all chitosan samples, including commercial one (Table 4). The crystal dimension decreased due to deacetylation because of the chemical treatment. Interestingly, crystallinity decreased in chitosan with respect to larval and adult chitin, but increased in pupal exuviae chitosan, suggesting that in pupal exuviae samples recrystallization can more extensively occur. Indeed, the alkaline treatment with 12 M NaOH and the successive steps, lead to a solubilization of chitosan in acetic acid solution. Hence, the crystallinity is generated after the final reprecipitation of the polymer. In the case of chitosan from adult chitin a higher order in the macromolecular chain is thus present, allowing a more extensive organization in crystals.

SEM

The surface morphologies of chitin and chitosan produced from H. illucens were observed by SEM and shown in Fig. 3a–c. First, chitin in the different sources, at 3000 × magnification, exhibited a structure with honeycomb-like arrangement, based on the repetition of square, pentagonal, and hexagonal units (Fig. 3a). Looking closer (magnifications 12,000–150,000×), the chitin samples showed significant surface differences (Fig. 3a,b). Various studies reported for chitin four different surface morphologies, such as (1) rough and dense surface without nano/microfibers and pores, (2) surface with combination of nano/microfibers and pores (the most common morphology), only fibrillar (3) and porous (4) surface^109,110. All unbleached and bleached chitin samples (Fig. 3b) consisted of scattered nanofibers with a diameter of about 30–50 nm. The surface complexity was found the highest for the adult chitin, whereas it becomes lower for pupal exuviae chitin and finally for larval chitin. Bleaching treatment had not significantly affected the chitin morphology of larvae and pupae exuviae whereas, for adult chitin, this process had removed some round particles from the fibrous arrangement. Chitins from larvae had a rough surface with broken fibers and an absence of pores that were present in chitins derived from exuviae and from adults. The surface of pupal exuviae chitins were denser than those of larvae, and not much porous. The micrometric morphology is evidently less regular than the one of adult chitin. Nanometric and micrometric holes peculiar to adult chitins revealed the presence of oriented nanofibers delimiting them (Fig. 3a(Ai–iii)). As reported by Purkayastha and Sarkar⁵⁶ and Soetemans et al.⁶⁴, the chitin derived from H. illucens adults showed different structural morphologies explained by the various extremities constituting the fly’s body. In agreement with the literature, the morphology of insect chitin can vary not only depending on the different species, and growth stage, but also on the genus and body part^89,90,97,109. Indeed, Kaya et al.^82,90,109 had described many different morphologies for adults of V. crabro, for a honeybee and between the females and males of some grasshopper species. Our results confirmed the presence of different chitin arrangement among different biomasses of H. illucens^{56,64,72,73,97}. According to its morphology, chitin can be used for different applications; particularly, chitin with a fibrillary surface is suitable for textile industry, while with a porous structure it can be used in tissue engineering and drug delivery^3,89.

All chitosan samples obtained from the larvae, pupal exuviae and adults of H. illucens (Fig. 3c) showed a rough but less fibrillated structure when compared to the respective chitin ones, demonstrating that deacetylation step altered the chitin structure, making it more homogeneous and less fibrillated. Indeed, the chemical chitin deacetylation deeply impacted its morphology. Chitosan nanofibers can be formed after reprecipitation of the solid polymer from acidic solution or being generated by modifying the previously existing chitin ones. Due to the higher morphologic complexity of adult chitin, we can hypothesize that the latter mechanism dominates in this case. On the contrary, the other samples, being less complex, can be more deeply modified resulting in a more effective formation of new nanofibrils. However, this fibrillated structure was more evident in unbleached than bleached samples, suggesting an influence of bleaching on the capacity of deacetylation to occur. As for chitin, the chitosan samples also had some pores on their surface. Due to the lack of studies on the surface morphology of chitosan produced from H. illucens, it was not possible to compare results of the present work with others in the literature. In an attempt to correlate the final physical–chemical properties of chitosan to the starting chitin structure, it is evident that the biomass of H. illucens is relevant. Adult chitin, showed a very high starting crystallinity and AD (linked to the high regularity of macromolecular structure), an intermediate value of viscosity average molecular weight (M_v) and a complex morphological structure resulted in a final chitosan showing the highest values of deacetylation degree (DD) and crystallinity. Hence, the morphological complexity of the sample is not significantly affected by these parameters. Pupal exuviae, showing the lowest crystallinity (linked to the lowest AD), an intermediate morphological complexity and a low M_v resulted in the lowest DD and intermediate crystallinity.

Finally, larval chitin, showing intermediate values of crystallinity, AD, the lowest morphologic complexity and the highest M_v, resulted in a chitosan with an intermediate DD and the lowest value of crystallinity. A low M_v in chitosan can be thus considered important for having a final high crystallinity, in agreement with the observations of Osorio-Madrazo et al.¹¹¹. A high starting AD seems another important element for having a high DD and crystallinity in the final chitosan. The bleaching, resulting in a strong decrease in M_v, determines an increase in crystallinity of the final chitosan. In general, despite different morphological structures were evidenced for the different biomasses, they did not extensively affect these important chitosan features, suggesting that the chemical attack due to deacetylation strongly modified the starting chitin morphological structure.

Potentiometric titration of chitosan

The DD of all chitosan samples produced from H. illucens is reported in Table 4. DD of all chitosan samples was around 90%, similarly to that of commercial chitosan, except for the unbleached sample from pupal exuviae (83%). DD reported by other authors for the chitosan produced from H. illucens was similar⁶⁴ or lower^57,66 than that measured in the present work, ranging from 40 to 90%. In most cases, chitosan from insects has a DD between 62 and 98%³², in accordance with the average DD reported for crustacean-derived chitosan (56–98%)¹¹². DD is an important parameter that influences different properties of chitosan, and it is dependent on the deacetylation conditions applied, in terms of temperature, reaction time and NaOH concentration; generally, higher temperatures can increase the DD¹¹³. A high DD enhances the antimicrobial activity of chitosan against certain bacterial species^114,115.

Viscosity-average molecular weight (M_v)

Molecular weight (Mw) for all chitosan samples was determined via viscometry and is reported in Table 4. M_v of all chitosan samples produced from H. illucens was much lower (from 21 to 92 kDa) than that of commercial chitosan (376 kDa). The M_v of chitosan samples from unbleached chitin was always higher than that of the respective bleached samples. These results confirm an effect of the bleaching treatment on the viscosity and M_v of the final chitosan^113,116. In particular, the bleaching reasonably decreased the Mw of the starting chitin inducing some scissions of polysaccharidic chains; probably the absence of catechol compounds making the polymer chains more accessible to partial hydrolysis too. Previous studies reported a M_v for insect chitosan in the range of 426–450 kDa^107,117. Considering the Mw determined by size exclusion chromatography techniques, insect chitosan ranges from 26 to 300 kDa³², whereas chitosan from crustaceans is reported to range from 100 to 1000 kDa^69,118. Much lower values of both M_v and Mw, between 3 and 10 kDa, have also been reported for insect chitosan, which are more similar to those obtained in the present work^89,95,119. Extremely low Mw can be due to a polysaccharide depolymerization caused by strong deacetylation conditions in terms of incubation time and temperature^32,69. Indeed, the same severe conditions can reduce the viscosity and thus M_v of the chitosan samples. We noticed that a lower M_v is generally linked to a higher crystallinity. Indeed, bleached chitosan samples had a lower M_v and higher CrI values compared to the respective unbleached samples. Mw can greatly affect physicochemical properties and biological activity of chitosan, often controlled by the macromolecular dimension. It is generally reported that chitosan with low Mw (< 150 kDa) has better antibacterial properties than high-Mw chitosan, since it can easier cross the cell wall of bacteria^120,121.

Chitosan film formation ability

All chitosan samples produced from H. illucens larvae, pupal exuviae and dead adults were able to form uniform homogeneous films. Figure 1c provides the photographic documentation. On the optical properties point of view, film of chitosan from adults (Fig. 1c(C,F) were the most different from all other samples, including commercial one: the unbleached chitosan film retained its dark brown colour, whereas the bleached one was much lighter but still more pigmented than the other bleached chitosan films.